Everyday Scientist

dylan, eat your heart out

Dylan at tenderbutton is always posting about his beautiful crystals. But I’ve one-uped him, because I did it the lazy way and with no usable results: I made crystals of my dyes by letting the waste beaker dry in the hood. I know, that’s a lab-safety no-no, but it was a mistake. And the results were cool!

Figure 1. The crystals under room lights.

Figure 2. The crystals under UV illumination. Note the heterogeneous fluorescence, a neato result of mixing a bunch of dyes and solvents together and letting evaporation do all the work.

Good Conduct

My lab is involved in several highly sensitive experiments that require complex data acquisition hardware. Much of our hardware is homemade, old, or inevitably unstable. As a result, we have to spend time fixing these systems, and that cuts into actual experiment time. To streamline the repair work and prevent future breakdowns, we have developed the flow chart shown in Figure 1.

Figure 1. Dishearteningly accurate.

I really like this figure. It’s truly a contender for best figure ever.

measuring total photons emitted

Good single-molecule fluorophores must meet several criteria. To name a few, the molecules must have high quantum yields, must be photostable, with a large absorption cross-section and a low yields into dark triplet states. (Ideally, a single-molecule fluorophore would have some inherent reporter function, but that’s a story for another day.) There are several popular long-lasting, bright fluorophores out there (e.g. rhodamines, terylenes, Cy3); my research involves developing and characterizing new compounds for single-molecule cellular imaging, which means quantifying the criteria I listed above.

One measure of photostability is the total number of photons each molecule emits before it irreversibly photobleaches (N_tot,emitted). Because photobleaching is a Poisson process that depends on the number of times the molecule reaches the excited state, N_tot,emitted for a fluorophore should not depend on the laser intensity (unless you illuminate beyond the saturation intensity): at a higher intensity, the molecules will absorb and emit photons at a higher rate, but photobleach in less time, yielding the same number of photons than at a lower intensity. So the N_tot,emitted of a particular fluorophore in a particular environment tells you how bright or long-lasting the fluorophore is; the higher the number, the better the fluorophore (all else the same).

I’ve measured the N_tot,emitted for a fluorophore on the bulk and single-molecule level—experiments which require different analysis. For the single-molecule measurement, I record movies of a wide-field illumination of several molecules well spaced in a polymer film. Then I integrate the counts from each molecule and plot the distribution from over 100 individual molecules. The distributions look like this:

The distributions are exponential characteristic of a Poisson process. From this distribution, I can determine the average N_tot,detected and convert this to N_tot,emitted using the detection efficiency of my setup (typically ~10% for epifluorescence).

The bulk measurement is a little less conceptually obvious. I use the same setup, but overdope the dye into the polymer film, so the intensity of the entire field of view versus time in the movie looks like this:

From the exponential fit, you can extract the average time before photobleaching for the molecules. This value, combined with the absorption cross section and the laser intensity, can give you the number of photons absorbed per molecule. Using the quantum yield, you can then calculate the N_tot,emitted.

So what’s my whole point here? Given that the two experiments I describe use different measurements and calculate N_tot,emitted using different parameters, I was really happy to see that the calculated values of N_tot,emitted were very close from bulk and single-molecule experiments when I compared them directly. That’s all.

[Update: You can find some of the relevant equations in this paper or the SI of this paper.]

Progress

Our 12 W Ar ion laser needed a tube change.* All it took was three weeks to convince Spectra Physics and two days with the service engineer. In the end, she’s a beaut’, Clark (see Figure 1).

Figure 1. Looking good, Billy Ray. Feeling good, Louis.

*The tube is changed every 6-12 months.

chilled water

The chilled water system at Stanford has been basically down for the last month and—now that the weather is getting ridiculously hot—we’re on the brink of the apocalypse. We can’t run our big lasers because they need to be cooled by the chilled water and the temperature in the offices and some of the labs it approaching hott with two “t”s. I mean, my office is in the basement and I’m sweating; the offices upstairs and in Mudd building are much warmer than outside (where the temperature is 90 F).

Thankfully, the most essential locations on campus (like the hospital) still get chilled water routed to them. Also, LINX, a for-profit cafeteria on the first floor of the BioX building, still is nice and cool. Thank god!

Check out this link for more cool (ha!) facts.

Lab Agriculture

I like to have a little life in lab. So, I plant various crops. In particular, cotton it a nice choice for any budding scientist, as shown in Figure 1. A nice pepper plant is also a good idea. The pepper is well suited for indoor growth, as shown in Figures 2 and 3.

Figure 1. God’s Q-tip.

Figure 2. Great stems.

Figure 3. Right after this picture was taken, my dog was run over by a tractor and a horse kicked my father.

lab-cleanup day

The Moerner lab had it’s (approximately) annual lab-cleanup day: We spent from 10am–3pm cleaning up the lab (with a break for lunch, of course). Here are some pics, including some before-and-after:

Figure 1. Look at everyone cleaning!

Figure 2. Unlabeled vial?!? I bet it’s dye.

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three lasers

In an older post, I talked about aligning multiple lasers on my setup. Now I have a cool picture of all of them (I used a little liquid nitrogen for scattering):

Those are lasers at 488, 532, and 633 nm.

dual-color viewer

How do you turn your grayscale CCD to a two-color camera? Filters and fun! Here’s a diagram looking down at the dual-viewer setup on my table:

Figure 1. Diagram of setup (viewed from above)

M1 and M2 are mirrors, F1 and F2 are long- or short-pass filters, and DC1 and DC2 are identical dichroics. DC1 reflects short wavelengths and pass long wavelengths, the filters clean up the two paths, the mirrors bring the paths back together, and DC2 combines the two channels. If the channels are offset a little, then short and long wavelengths are split into two copies of the image onto the CCD.
Here’s a pic of the setup:

Figure 2. Picture of the setup with channels drawn

In other words, the dichroics split the green light off the output and move it to a different region of the CCD. You can recombine the two copies and add color using ImageJ, like this:

Figure 3. The right side is an overlay of the red and green channels in false color

lab inspections

Stanford and the county are very careful about lab safety and waste disposal. Maybe it’s just California regulation, but sometimes it goes a little over the top. My new favorite requirement: Dalbir, who is our department’s Safety Compliance Officer, tells us that we need secondary containment (i.e. a bucket) for our propane torch. I think that’s pretty stupid: I’m pretty sure propane isn’t liquid at room temperature and pressure…

comparing gaussian structures

Now, there’s probably already a program out there that will compare the optimized structures of quantum-chemistry calculations (e.g. Gaussian). But I couldn’t find one, and I wanted control over how to compare the structures (for instance, I wanted to be able to ignore the errors in the dihedrals for light atoms). So I wrote a MatLab M-file that outputs the absolute differences in the bond lengths, angles, and dihedrals between two structures. It took me a while to get what I wanted out of Molden and make MatLab spit out the right set of numbers, but I finally got it to do what I wanted (I think).

It’s not perfect, I’m pretty happy with it. It just imports the variables from a Molden z-matrix and takes the absolute difference in them, sorting my type. Then I just throw those outputs into SigmaPlot or Excel (ew!) or whatever and do the error analysis I want. For instance, I remove the dihedrals that don’t really matter (some light atoms and some artificially inflated dihedral errors) and just look at the mean and standard deviation for different method/basis set combinations.

I dunno, not essential. But I find it pretty handy. If you want to check it out or try it, I’ll post it below. Let me know if you find errors or if you find ways to make it better.

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easy as 1, 2.1, 2.2, 2.3, 2.4, 3.1, 3.2, 3.3, 3.4

Labs at Stanford are supposed to have all these safety posters everywhere: we’re super cautious about lab safety. Part of this has to do with the hyper-regulation in California. Here’s one of my favorite posters:

It’s so easy now! Really, the best part is that there are even bulleted subsections under sections under numbers. That’s more than simple: it’s complex!

single molecules with a digital camera!

These are single fluorescent molecules imaged using a microscope and a hand-held consumer digital SLR camera (i.e. Nikon D90):

I think that’s pretty impressive. Usually, we use expensive, cooled CCD cameras which are very sensitive and designed for scientific imaging. Here, I used a (cheaper) conventional digital camera and even got a color image. This is possible in part because this fluorophore (one of the Moerner/Twieg labs’ DCDHF dyes) is super bright and long-lived!

f=e^(very complicated)

Dealing with engrained technology can be a little frustrating. Many years ago, a grad student wrote a program in LabView to fit Gaussians and exponentials. This is a simple problem, and there are many fine programs for dealing with numerical analysis. The problem with LabView is that it isn’t one of them. As shown in Figure 1, it gets a little complicated.

To be fair, it does work. It fits mighty nicely after it is debugged and running, as shown in Figure 2. As my co-worker says, “When it works it works well.”

Figure 2. A good fit.

3-d surface

A fellow labmember showed me how to use MatLab to make a 3-dimensional surface from an image of single fluorescent molecules, where z is the intensity. Here’s a taste. I converted this:

into this:

The funny thing is that the pretty image is only that: it actually contains less information than the 2-d grayscale image, becuase it needed smoothing and convolving with Gaussians to make it look so nice. But that kinda shit brings in the big $$.